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We examined the effect of delivery modality on the survival, localization, and functional effects of exogenously administered embryonic stem cells (ESCs) or endothelial cells derived from them (ESC-ECs) in the ischemic hindlimb.
Murine ESCs or ESC-ECs were stably transduced with a construct for bioluminescence imaging (BLI) and fluorescent detection. In a syngeneic murine model of limb ischemia, ESCs or ESC-ECs were delivered by intramuscular (IM), intra-femoral artery (IA), or intra-femoral vein (IV) injections (n=5 in each group). For 2 weeks, cell survival and localization were tracked by BLI and confirmed by immunohistochemistry, and functional improvement was assessed by laser Doppler perfusion. BLI revealed that ESCs localized to the ischemic limb after IM or IA, but not after intrafemoral vein administration. Regardless of the route of administration, ESCs were detected outside of the hindlimb circulation in the spleen or lungs. ESCs did not improve limb perfusion and generated teratomas. In contrast, ESC-ECs delivered by all 3 modalities localized to the ischemic limb, as assessed by BLI. Most surprisingly, ESC-EC injected IV eventually localize to the ischemic limb after initially lodging in the pulmonary circulation. Immunohistochemical studies confirmed the engraftment of ESC-ECs into the limb vasculature after 2 weeks. Notably, ESC-ECs were not detected in the spleen or lungs after 2 weeks, regardless of route of administration. Furthermore, ESC-ECs significantly improved limb perfusion and neovascularization, when compared to the parental ESCs or the vehicle control group.
In contrast to parental ESCs, ESC-ECs preferentially localized in the ischemic hindlimb by IA, IM, and IV delivery. ESC-ECs engrafted into the ischemic microvasculature, enhanced neovascularization, and improved limb perfusion.
Peripheral arterial disease (PAD) affects about 10 million Americans, is typically due to atherosclerotic arterial occlusive disease of the limbs, and may be manifested as intermittent claudication, ischemic ulcerations, or even limb-threatening gangrene.1 Therapeutic angiogenesis is under investigation as a novel approach to the treatment of this condition. Local administration of angiogenic genes or proteins have shown promise in preclinical studies, but in randomized clinical trials, improvement (eg. by treadmill exercise capacity) has been modest and transient at best. An approach that might have a greater and more persistent effect would be to administer cells that can produce angiogenic cytokines and/or participate in vascular regeneration. Pre-clinical studies revealed the existence of bone-marrow derived mononuclear cells (BM MNCs) that could incorporate into the microvasculature, and/or produce angiogenic cytokines that increase microvascular density and perfusion in the setting of ischemia.2, 3 Subsequently, several well-controlled clinical studies have provided evidence that local administration of bone-marrow derived mononuclear cells (BM MNCs) can reduce the extent of myocardial infarction, improve perfusion of the ischemic myocardium.4, 5 and improve ventricular function (BOOST and REPAIR-AMI).6-9 With respect to PAD, BM MNCs have shown encouraging results in small clinical trials.10, 11
The mechanisms whereby adult stem cells improve myocardial or peripheral perfusion in patients with ischemic disease are not fully understood. Furthermore, the BM MNCs that promote angiogenesis are incompletely characterized. But the major limitation to the effective clinical application of adult stem cell-based therapy is that in patients with cardiovascular disease, the circulating BM MNCs that promote angiogenesis are reduced in number and function.12-14 Accordingly, an alternative approach would be to generate sufficient quantities of functional endothelial cells from pluripotent stem cells.
Previous studies have suggested that administration of ESC-ECs may enhance perfusion in the murine ischemic hindlimb.15-18 In the current study, we extend these observations by employing molecular imaging, genetic markers and laser Doppler blood perfusion (LDBP), to gain new insights into the localization, survival, and functional improvement induced by ESC-ECs.
Murine ESCs (D3, ATCC, Manassus, VA) of the SV129 strain were cultured on mitotically inactivated mouse embryonic fibroblasts as previously described.19 ESCs stably transduced with a double fusion reporter construct containing the ubiquitin promoter driving firefly luciferase (fluc) and monomeric red fluorescence protein (mRFP) were cultured under the same conditions. The double fusion construct was previously shown neither to interfere with ESC phenotype nor differentiation capacity.20
Endothelial differentiation of ESCs was carried out using the suspension culture approach with modifications.21 To initiate differentiation, ESCs were cultured in ultra-low non-adhesive dishes to form embryoid body (EB) aggregates in a differentiation media that consisted of Alpha Minimum Eagle's Media, 10% FBS, 1% penicillin/streptomycin, and 0.05 mM β-mercaptoethanol (Sigma, St Louis, MO). After 4 days of suspension culture, the EBs were reattached onto 0.2% gelatin-coated dishes and cultured in differentiation media. After 3 weeks of differentiation, the cells were purified by fluorescence activated cell sorting (FACS) using anti-mouse VE-cadherin Ab (BD Biosciences, Bedford, CA). To purify ESC-ECs, single cell suspensions were obtained by treatment with cell dissociation buffer (Invitrogen) and then passed through a 70-μm cell strainer (BD Biosciences). Cells were incubated with rat anti-mouse VE-cadherin Ab for 20 minutes, followed by allophycocyanin (APC)-conjugated anti-rat IgG2a (BD Biosciences) Ab for 20 minutes. Isotype-matched Abs served as negative controls (Millipore). The VE-cadherin-expressing cells were isolated using the FACStar (BD Biosciences) cell sorter and maintained in differentiation media supplemented with 50 ng/ml vascular endothelial growth factor (VEGF).
Immunofluorescence staining, acetylated low-density lipoprotein (Ac-LDL, Invitrogen) uptake assay, and matrigel tube-like formation assays were used to verify the phenotype of ESC-ECs. Immunofluorescence staining of ESC-ECs was carried out using EC markers, namely VE-cadherin, von Willebrand factor (VWF, Abcam), and endothelial nitric oxide synthase (eNOS, BD), according to established methods.22 Briefly, samples were fixed in 4% paraformaldehyde, permeabilized with 0.5% Triton X-100, and pretreated with 1% bovine serum albumin (BSA). After incubation with primary Abs, alexafluor-488-conjugated secondary Abs (Invitrogen) were applied. Cell nuclei were stained by Hoechst 33342 (Invitrogen). Uptake of Ac-LDL was assessed by incubating ESC-ECs with 5 μg/ml alexafluor-594-conjugated Ac-LDL (Invitrogen) for 5 hours before detection by fluorescence microscopy.20 The formation of endothelial tube-like structures was assessed by growing cells for 24 hours on growth factor-reduced matrigel (BD Biosciences).20
For non-invasive tracking in vivo, the purified ESC-ECs were transduced with a lentiviral vector carrying an ubiquitin promoter driving fluc and enhanced green fluorescence protein (LV–pUb–Fluc–GFP) as described previously.19 The transduced cells were isolated using FACS by GFP positive expression and maintained in differentiation media containing 50 ng/ml VEGF. To determine the correlation between cell density and fluc activity ex vivo, cells were incubated with reporter probe D-luciferin (150 μg/ml), and bioluminescence imaging (BLI) was performed with the In Vivo Imaging System 200 system (IVIS 200; Caliper Life Sciences, Hopkinton, MA). The BLI intensity was expressed in units of photons/cm2/s/steradian (p/cm2/s/sr). In addition, to verify GFP expression and the maintenance of EC phenotype, the transduced cells were analyzed by FACS and immunohistochemistry for dual expression of GFP and VE-cadherin.
To assess the survival and localization, and potential therapeutic potential of ESCs, we used a murine model of HLI, according to our previous reports.23, 24 Briefly, unilateral HLI was induced by ligating the femoral artery of 8–10 months-old female syngeneic mice. Aged mice were utilized because young animals more easily recover in the absence of therapeutic intervention.25 The mice were randomized into several groups, each receiving 106 transduced ESCs by intra-femoral artery (IA), intra-femoral vein (IV), or gastrocnemius intramuscular (IM) routes (n=5 each group). Additional control animals underwent HLI induction only. To test the survival, localization, and therapeutic potential of ESC-ECs, a similar study was carried out in which 5×104 transduced VE-cadherin-expressing ESC-ECs were delivered by IA or IM delivery, and 5×105 ESC-ECs were delivered by IV route (n=5 each group). Animal studies were approved by the Administrative Panel on Laboratory Animal Care in Stanford University.
At specified time points up through 2 weeks, BLI was performed on animals after injecting D-luciferin (375 mg/kg body weight) reporter probe into the peritoneum, as described previously.26 Using the Living Image 3.0 software (Caliper Life Sciences), regions of interest (ROIs) were drawn over the ischemic hindlimb or the lung, and BLI intensity was quantified as described.26, 27 To determine the threshold for positive BLI signal, animals without cell injection were imaged and quantified accordingly.
Blood flow to the ischemic or normal (non-ischemic) hindlimb was assessed using a PeriScan PIM3 laser Doppler system (Perimed AB, Sweden) as described previously.23, 28 Each animal was pre-warmed to 37°C core temperature, and hindlimb blood flow measured pre- and post-operatively on day 0, and again on days 4, 7, and 14. ROIs of the ischemic or non-ischemic hindlimb were drawn in a standard fashion. The level of perfusion in the ischemic and normal hindlimbs was quantified using the mean pixel value within the ROI, and the relative changes in hindlimb blood flow were expressed as the ratio of the left (ischemic) over right (normal) LDBP.
After 2 weeks, the mice were euthanized, and the hindlimbs were snap frozen in OCT compound (Sakura Finetek, Japan) and cryosectioned for histological and immunofluorescence assessment. Representative tissue sections were processed for routine hematoxylin and eosin (H&E) 29 staining and assessed by a pathologist. Images were captured using a Nikon Eclipse E1000M microscope (Nikon, Burlingame, CA) for the evidence of teratoma formation. Incorporation of exogenous cells into the microvasculature of the ischemic tissue was assessed using double immunofluorescence staining for CD31 (BD Biosciences) and GFP. At least 4 tissue sections were assessed for each animal, and 5 high-powered images were photographed for each tissue section with a SPOT RT color camera (Diagnostic Instruments, Sterling Heights, MI). Capillary density was quantified using Image J software (NIH, Bethesda, MA) by counting the number of CD31+ capillaries and averaging among the 4 sections for each sample, according to previous studies.17, 30, 31 The contribution to capillary formation by the transplanted cells was quantified by counting the number of capillaries expressing both GFP and CD31. Total and GFP+ capillary densities were expressed as number of capillaries/mm2.
To determine the localization of transduced ESCs or ESC-ECs outside of the hindlimb in vivo, the spleen and lungs were harvested, and genomic DNA was isolated using the DNEasy Kit (Qiagen) to detect fluc expression, normalized to GAPDH levels. The primers consisted of fluc forward primer, ATCTACTGGTCTGCCTAAAG, reverse primer, CAGCTCTTCTTCAAATCTATAC; and GAPDH forward primer, TTCACCACCATGGAGAAGGC, and reverse primer, GGCATGGACTGTGGTCATGA, based on previous studies.32 The PCR products were separated on 1% agarose gel electrophoresis and imaged using the Alpha Imager 3400 (Alpha Innotech). To determine the threshold of cells to detect positive fluc gene expression, varying numbers of cells were combined with spleen or lung tissue and assayed by PCR.
All data are reported as mean ± standard deviation. Statistical analysis between multiple groups at each time point was performed by analysis of variance (ANOVA) with Holm's adjustment. Repeated measurements of samples over time were analyzed by repeated measures ANOVA with Holm's adjustment. Statistical significance was accepted at P<0.05.
We delivered cells to the ischemic limb by IA, IV, or IM modalities and tracked by BLI for 2 weeks (Figure 1). Previously we have shown that the BLI signal directly correlates with cell number for ESCs stably transduced with a construct optimized for non-invasive tracking.20 Cells delivered by the IA route were primarily localized in the ischemic limb (Figure 1A). After the initial administration, the intensity of the BLI signal increased and was significantly higher at day 14 by comparison to day 0 (3.1±1.2×106 vs 7.5±4.9×104 p/s/cm2/sr; P<0.005). Although BLI signal in the lung was not greater than threshold at day 0, the signal exceeded the threshold value by day 14 (4.6±8.7×104 p/s/cm2/sr) (Figure 1F). In the group receiving cells by IM delivery, the kinetics of cell survival and localization were similar (Figure 1C). By contrast, in the group administered ESCs by the IV route, a BLI signal never exceeded threshold values in the ischemic hindlimb at any time point, but a strong signal was detected in the lungs (Figure 1B). On day 0, the signal intensity in the lungs was significantly higher in the IV group compared to other groups (2.1±1.2×105 vs 2.6±0.9×103 vs. 1.7±0.6×103 p/s/cm2/sr; IV vs IA vs IM; P<0.005). The BLI intensity in the lungs declined from day 0 to day 3, but increased by day 14 (Figure 1F), which suggested initial cell loss followed by proliferation of ESCs in the lungs.
To verify the localization of transduced ESCs outside of the hindlimb, we assessed the expression of fluc in lung and spleen by isolating genomic DNA from these tissues at the time of euthanasia. Figure 1E shows fluc and GAPDH expression by PCR for each animal (n=5 in each group). We observed positive bands for at least one animal in each cell-treated group in either the spleen or lungs, whereas the control animals showed no fluc expression (Figure 1E). These results substantiated the bioluminescence data showing the persistence of cells in the lungs after 14 days. We also characterized the sensitivity of PCR for detecting fluc expression by isolating genomic DNA from tissue samples containing various cell numbers. Based on the sensitivity assay, the threshold for PCR detection was ≥104 cells in the spleen and ≥103 cells in the lung (Supplemental Figure I).
Postmortem histological assessment of the hindlimb tissue sections at 2 weeks after ESC delivery revealed early signs of teratoma formation. As shown in Supplemental Figure II, H&E-stained tissue sections revealed proliferation of the engrafted cells associated with mitotic activity and apoptotic figures. Areas of necrosis alternating with viable cells that focally formed ducts and cyst-like structures were observed. Supplemental Figure IIA shows well-developed columnar epithelial cells surrounding a central lumen forming a duct-like structure, whereas Supplemental Figure IIB–C shows early formation of cystic structures, lined by cells resembling keratinized squamous epithelium. Thus ESCs could survive and remain generally localized in the ischemic limb by IA or IM delivery, but evidence of cell escape to the lungs or spleen was observed. Furthermore, early teratoma formation was observed in the ischemic hindlimb in the IM or IA ESC groups.
We next determined the effect of ESC delivery on hindlimb perfusion by LDBP assessment. As shown in Figure 2A–B, there was no improvement in hindlimb mean perfusion ratio (ischemic/control hindlimb) in any cell treatment group on day 14 (IA, 0.49±0.18; IV, 0.57±0.20; IM, 0.63±0.22), in comparison to the control group (0.57±0.17). Indeed, hindlimb perfusion was reduced on day 3 in the IA (0.33±0.07) and IV (0.39±0.11) groups, in comparison to the control group (0.59±0.17; P<0.005). These results suggest that delivery of ESCs by any of the 3 modalities does not enhance (and may even delay) the recovery of hindlimb perfusion, and serves as a useful control for the following studies.
ESC-ECs were purified by FACS based on positive VE-cadherin expression after 3 weeks of differentiation. By comparison to published protocols that purify the cells after 2 weeks of differentiation, we observed improvement in yields by prolonging differentiation time from 2 to 3 weeks prior to purification. In our hands, this modification resulted in yields as high as 50%, but with typical yields of 10-30% (Figure 3A). Re-analysis of the sorted cells demonstrates that >97% of the cells were indeed VE-cadherin+ cells, demonstrating the purity of the ESC-ECs (Figure 3A). The purified cells were expanded and characterized by immunofluorescence staining for EC markers such as VE-cadherin, VWF, and eNOS (Figure 3B). In addition, the ESC-ECs could efficiently incorporate Ac-LDL (Figure 3B) and form tube-like structures in matrigel (Figure 3C). By comparison, the parental ESCs could not form tube-like structures (Figure 3C). Together, these studies confirmed the endothelial phenotype of the ESC-ECs.
In order to track the localization and retention of ESC-ECs in vivo, we stably transduced the cells with a double fusion reporter construct (Supplemental Figure IIIA) and isolated the transduced cells by FACS based on dual expression of GFP and VE-cadherin. Over 80% of the ESC-ECs manifested GFP and VE-cadherin expression (Supplemental Figure IIIB). Furthermore, as shown in Supplemental Figure IIIC, there was a strong correlation between BLI signal and cell number, indicating that BLI could be used for quantification. After transduction, the ESC-ECs expressed GFP and maintained their endothelial phenotype, based on double expression of GFP and VE-cadherin (Supplemental Figure IIID).
In comparison to the patterns of ESC localization, we observed salient differences using the ESC-ECs (Figure 4A–F). Immediately after cell delivery, BLI signals were 7.5±5.2×104 p/s/cm2/sr for the IA group and 1.5±2.5×105 p/s/cm2/sr in the IM group, with no observable signal in the lungs for either group. After 14 days, the cells remained localized to the hindlimb in the IA (2.4±5.4×105 p/s/cm2/sr) and IM (4.6±7.0×105 p/s/cm2/sr) groups, without a positive signal in the lungs (Figures 4D, 4F). The BLI data was supported by PCR results, as fluc expression could not be detected in the lungs or spleen (Figure 4E). The ESC-ECs delivered by IV injection showed minimal BLI signal in the ischemic hindlimb on day 0 (1.9±1.1×104 p/s/cm2/sr). Intriguingly, a BLI signal became appreciable by day 3 (5.1±10.5×104 p/s/cm2/sr) and reached levels that were similar to that of the other ESC-EC groups by day 14 (1.4±3.0×105 p/s/cm2/sr) (Figure 4B, 4D). Cellular localization in the lungs was significantly higher for the IV group at day 0 (2.3±1.5×106 vs. 5.7±2.8×103 vs. 7.3±4.6×103 p/s/cm2/sr; IV vs. IA vs. IM groups; P<0.005). However, BLI signal diminished in the lungs in the IV group with time until it was no longer detectable at day 14 (Figure 4F). PCR results also confirmed the absence of fluc expression in the lungs and spleen by day 14 (Figure 4E). Notably, when ESCs and ESC-ECs were administered IV, each cell type was found primarily in the lung on day 0. However, only the ESC-EC signal disappeared from the lung, in association with appearance of the signal in the ischemic hindlimb. The differences in BLI localization of ESC-ECs and the parental ESCs were confirmed by PCR of tissue samples derived from the spleen and lung of experimental animals at 2 weeks.
Immunofluorescence staining of tissue sections verified the localization of ESC-ECs in the ischemic hindlimb at 2 weeks after delivery. The transplanted cells were identified based on the co-localization of GFP and CD31 expression. For all 3 groups treated with ESC-ECs, the majority of visualized cells were found in association with capillaries in the ischemic limb and less frequently in association with larger vessels (Figure 5A). Furthermore, ESC-ECs in the IM group were localized only in the gastrocnemius near the sites of cell injection, whereas in the IV and IA treatment groups, the ESC-ECs were found throughout the hindlimb spanning the thigh and gastrocnemius muscles (Supplementary Figure IV).
We next determined whether ESC-EC delivery could provide any improvement in hindlimb angiogenesis and perfusion. The capillary density attributed by ESC-ECs were significantly higher in the IA (98±14 capillaries/mm2) and IV (71±23 capillaries/mm2) treatment groups, compared to the IM (37±14 capillaries/mm2) group, based on colocalization of GFP and CD31 (P<0.03) (Figure 5A and 5B). Total capillary density analysis also demonstrated significantly increased density in the IA (1120 ±133 capillaries/mm2) and IV (1136±266 capillaries/mm2) treatment groups, when compared to the control (868±127 capillaries/mm2) group (P<0.05) or IM (757±52 capillaries/mm2) group (P<0.003) (Figure 5C-D). The IM group did not show any statistically significant improvement when compared to the control group (Figure 5C-D). The improvement in angiogenesis was further supported by laser Doppler blood perfusion analysis. As shown in Figure 6, there was a greater improvement in hindlimb mean perfusion ratio for the IV (0.82±0.13) and IA (0.76±0.07) groups at day 14, in comparison to the control group (0.57±0.17; P<0.005). The increase in perfusion ratio for the IM group was not significantly different from control. These results demonstrated that intravascular delivery of ESC-ECs could significantly improve limb perfusion.
The salient findings of this study are that 1) in a murine model of hindlimb ischemia, syngeneic ESC-ECs preferentially localize in the ischemic hindlimb; 2) the ESC-ECs appear to maintain their EC lineage and incorporate into the microvasculature; and 3) the localization of ESC-ECs in the ischemic hindlimb is associated with a greater improvement in limb perfusion. By contrast, the parental ESCs do not preferentially localize to the ischemic hindlimb and do not improve perfusion (and in fact, can differentiate into non-vascular cells). This is the first study to combine molecular imaging, genetic markers, and LDBP to gain new insights into homing, incorporation, and survival of ESC-EC in hindlimb ischemia. Furthermore, this is the first study to demonstrate that ESC-ECs administered systemically have the capacity to localize to a site of ischemia.
Localization of cells to the ischemic tissue after IV administration was assessed by BLI. After IV administration, the signal persisted in the lung to day 14 in the animals treated with ESCs. By contrast, in the group that received ESC-ECs, the signal in the lungs diminished to undetectable levels by day 14, while the signal increased in the ischemic hindlimb. These observations are consistent with a re-distribution of the ESC-ECs from the lung microvasculature and to the ischemic hindlimb. This hypothesis was supported by immunohistochemistry and fluc expression studies of the ischemic hindlimb and lung. Previous work have implicated roles for VEGF, the chemokine (C-X-C motif) receptor 4 (CXCR4) and its ligand, stromal-derived factor (SDF-1) in this homing phenomenon.33, 34 Another explanation for the kinetics of ESC-EC signal could be a depletion of cells in the lung, concomitant with proliferation in the hindlimb. This explanation is unlikely, as evidence of cell proliferation was not as pronounced in animals receiving ESC-ECs by the IA or IM route. The approach of molecular imaging and genetic markers as used in the current investigation will facilitate further elucidation of the proclivity for ESC-ECs to localize to the ischemic region.
Ours is the first study to show that ESC-ECs administered systemically can, over a period of several days, localize to a site of ischemia and improve perfusion. In this way, ESC-ECs recapitulate the homing behavior of bone marrow derived endothelial progenitor cells. Previous studies using the murine ischemic hindlimb model have utilized IM or IA delivery. Two groups have shown that in young immunodeficient mice, the IM administration of human ESC-ECs improve blood perfusion after 4 weeks.15,16 Human ESC-ECs have also been administered by IA injection to young nude mice, and shown to incorporate in the vasculature and improve angiogenesis and perfusion with increases in capillary density and blood perfusion.17, 18 A limitation of these studies is that they used young immunodeficient animals. By contrast, we used older immunocompetent mice. Our aged mice (8–10 months old), like patients with PAD, have impaired vascular function and regenerative capacity by comparison to young animals.25 Furthermore, by incorporating syngeneic ESC-murine ECs, our study avoids the confounding variables of immunodeficiency (which may have unanticipated effects on bone-marrow derived cells that participate in angiogenesis), and inter-species differences. Finally, our model anticipates vascular regeneration using therapeutic cells derived from autologous stem cells.
This is the first report utilizing molecular imaging approaches to non-invasively track the survival and localization of ESC-ECs into the ischemic hindlimb over time. Using molecular imaging to track cell retention and localization, along with LDP for functional assessment, this body of work provides a foundation for understanding the role of delivery modality on therapeutic efficacy. Our studies suggest that systemic delivery of ESC-ECs may be superior to localized IM delivery, and achieve similar efficacy as intra-arterial administration.
We used double fusion reporter constructs to enable dual tracking by BLI as well as by a fluorescent protein. We previously showed that this lentiviral construct did not interfere with ESC phenotype and differentiation capability19, 20 and our results suggest that the transduced ESC-ECS retained their endothelial phenotype after lentiviral transduction. BLI is a sensitive and accurate method for tracking cells in vivo with as little as 500 cells.19 Traditional methods of tracking transplanted cells using fluorescent reporter genes require intensive histological assessment to identify the transplanted cells and are difficult to quantify.
Using BLI, it is possible to characterize the survival kinetics of ESC-ECs as well as ESCs in the ischemic hindlimb. In other ischemic tissues such as the infarcted myocardium, we previously showed that ESC-ECs engraft but undergo cell loss, ultimately leading to <10% survival after 2 weeks.20 In contrast, the current study suggests no significant loss of ESC-ECs over time, suggesting that the disease model may affect the survival of transplanted ESC-ECs. Long-term studies to track cell survival and localization would be interesting and warranted. BLI of ESCs revealed an exponential increase in cell number after 2 weeks with all three modalities of cell delivery. The behavior of ESCs in our model is consistent with previous observations of the temporal kinetics of ESC proliferation in ischemic tissues.20 Not unexpectedly, early teratoma formation was observed in the animals treated with ESCs.20, 35 On the other hand, there appeared to be no signs of teratoma formation after delivery of ESC-ECs after 2 weeks, although longer time points would be needed to preclude the possibility of teratoma formation.
In conclusion, we used molecular imaging and genetic markers to track the localization of ESCs or ESC-ECs in the ischemic hindlimb, and to demonstrate the role of delivery modality on ESC or ESC-EC survival and therapeutic efficacy. Using transduced cells optimized for non-invasive imaging, we found that, in comparison to the parental ESCs, ESC-ECs preferentially localize in the ischemic hindlimb. Post-mortem immunofluorescence staining confirmed the engraftment of ESC-ECs in the microvasculature of the ischemic hindlimb. Furthermore, intravascular delivery of ESC-ECs is associated with enhancement of limb perfusion. This study provides a foundation for non-invasive monitoring of the localization and survival of therapeutic cells, and translational application in the treatment of PAD.
This study was supported by grants to JPC from the National Institutes of Health (U01HL100397, RC2HL103400, R01CA098303, R21HL085743, 1K12HL087746, 1P50HL083800), the California Tobacco Related Disease Research Program of the University of California (18XT-0098), the California Institute for Regenerative Medicine (RS1-00183), American Heart Association (0970036N), and the Stanford Cardiovascular Institute; and to JCW from the National Institutes of Health (R21HL091453). N.H. was supported by a fellowship from the American Heart Association.
Disclosures: No conflicts declared.